Yup, wow, this is a big experiment! Some tips that might help you...

1) It does not matter how you number/label samples, as long as you know what they are, and the person who comes after you knows what they are! Writing out tons of info onto tubes is a waste of time (IMHO). I number things, and make sure the box they're stored in refers to the Excel sheet where the numbers are listed against detailed information about each sample.

2) 100mM sounds like an unlikely concentration for primers - this would be extremely high. The most common concentration to resuspend stock primers is 100uM. If you want to 'sanity check' the concentration of the frozen primers, run an aliquot (maybe 5uL) on a high percentage agarose gel (2-3%) alongside a primer of known concentration - at least this should tell you if the concentration is different by an order of magnitude or something.

3) If those frozen primers are in original tubes from the company that synthesised them, and they were ordered through an account you have access to, you might be able to look up documentation. I know IDT keeps info / spec sheets available online for several years after the order was placed.

4) For initial testing of primers and optimising assays, your two best friends are practice samples and pooled samples. Whenever possible, I like to start with practice samples: these are as similar to my experimental samples as I can possibly get (same species, sex, strain, approximate age, tissue type, extraction method, etc etc), but have no experimental value. These are usually fairly easy to get. Does someone else in the building work on rats of the same strain and age, but they use a different organ to you? Ask if you can be present when they are euthanising animals to collect practice tissue. Does your animal facility cull animals that are excess, wrong genotype etc? Ask if you can have the cadavers. In most places, tissue sharing of this kind is permissible without specific animal ethics approval; if you do need approval, it should be trivially easy to get. If you have a large amount of practice tissue / practice samples, you can test things as many times as you need to without any anxiety that you will run out of precious samples. In my opinion, this reduces stress, and reduces the likelihood that you will have to proceed with a sub-optimal method or publish a questionable result simply because you ran out of sample to do it again properly.

If it's really not possible to get practice samples that are similar enough to your real ones, or if you're in the final stages of optimising, that's when you can use pooled samples. To do this, you can take a little RNA or cDNA (whichever you currently have) from each sample and combine them together in one tube. Then use this pooled cDNA as the substrate for test PCRs. If you did this once per group, you would have n=8 instead of n=48, and you would use only 1/6th as much material from any one sample as if you tested on all 48 samples individually. This reduces both sample and reagent consumption during workup experiments.

Also just checking - are you aware that you will have to do a separate cDNA synthesis to test miRNA levels?

Dear Laura Leighton! Many thanks for the detailed answer with so precious info. I suspected there are algorithms)))

1) I ponder about sharing the Excel sheet among colleagues involved in the project.

2) Nice idea! Thanks.

3) Unfortunately there are only aliquotes in microtubes.

4) What a beautiful method. Will definitely use it.

5) Yes, I am aware of a long miRNA protocol...

Thanks again!

(1) Organize your samples, including any positive and negative controls. You should at minimum have a negative "no template" control (just your primers and master mix) so you can figure out which genes aren't expressed in any sample at all and just show primer dimer signal. Positive controls are also really nice to have. In the past, I have purchased commercially made cDNA as my positive control to control for tissue-specific lack of expression, or cDNA synthesis failure. Pooled samples can also be used as semi-positive controls in case there's sample-specific lack of expression.

(2) Organize your plate layouts on Excel. Color-code 8x12 cells (assuming you are using a 96 well plate) and label A-H on one side, 1-12 on the other side. Know exactly where your sample and gene combinations will be.

(3) Group & separate your samples so that the downstream analysis will be correct. If you are doing ddCt analysis to compare gene expression, then pro tip, you just need to have ALL your samples *OR* ALL your genes (or a whole group of genes) on one plate. Obviously it's too many samples to have all variables on one plate, but if you pick one or the other, the ddCt will work out and cancel out plate-to-plate batch effects better that way. I suggest trying all your samples on each plate, and setting up multiple plates to get through all genes. That way you can redo a plate if needed, without messing up your experiment.

(4) I recommend treating the protein coding genes, long noncoding genes, and miRNA as 3 separate experiments. The long noncoding are likely to have weaker expression so you may need more cDNA. The miRNAs are the length of a primer so you need to increase the length during the cDNA step so you are able to PCR-amplify them. If you put all your samples on one plate, you can space out the genes on different plates. You can also use different reference genes for each of the 3 groups. It's recommended that your reference genes be roughly similar expression to your experimental genes, so you don't want a 9000 transcripts per million (TPM) reference and a 10 TPM long noncoding gene. Usually the references are higher expressed to make sure you get results from each sample, but a more reasonable difference would be a 100 TPM reference for a 10 TPM experimental gene.

(5) If there is any published RNA-seq related to your work, use it to check that your reference genes are ok (not significant, very high p-value and low fold-change). Make sure you pick reference genes that are not differentially expressed between your study groups. You can also check TPM values (or FPKM values) from RNA-seq results to estimate gene expression levels.

(6) When pipetting everything, open a new tip box and take advantage of the 8x12 layout to match your plate. Keep the order matching. Don't rely on memory.

(7) Lastly, if your lab has the funding, consider nanofluidic qPCR methods. You need less sample that way and you can run 96x96 well plates. You load your primers on one side, samples on the other side, and run a chip that way. See if your institution has a genomics core and if they have the machines for running nanofluidic qPCR chips. I have used Fluidigm in the past and it works well.

Tania L Gonzalez thanks for the tips!

1/Please, explain what you mean by the miRNAs are the length of a primer so you need to increase the length during the cDNA step so you are able to PCR-amplify them? What is the increasing length - during the reverse transcription step?

2/ What mean to take advantage of the 8x12 layout to match your plate? Usually, I place pipette tips from a newly opened huge pack in a special box before work.

P. S. Is there any free calculator for concentrations? I`m still confused while calculating the amount of primers/cDNA for PCR reaction volumes(

Anna Zhukovska , you're welcome!

1/I assume you are using mature miRNAs which tend to be about 20-24 bases long. Primers are also about that size, so you can't design two primers to bind either end of the miRNA (they would just dimerize and amplify each other). You need to make the cDNA sequence longer in two reactions: a polyadenylation reaction that adds a short poly-A tail to the miRNAs, then a reverse transcription reaction with primers that adds extra sequence to make the cDNA even longer.

I have run RT-qPCR on miRNAs two ways. Both required a polyadenylation reaction, but the cDNA synthesis step was different. The do-it-yourself way is to design your own cDNA primers. I used the method in Busk 2014, "A tool for design of primers for microRNA-specific quantitative RT-qPCR", doi: 10.1186/1471-2105-15-29. Busk has a script you can run. Input your miRNA sequences, and Busk's script gives you primers for cDNA synthesis. Try with his example first to make sure that it works, then make primers for your own miRNAs. The upside is that you can run a regular qPCR reaction after this. The downside is that you need to make a different cDNA synthesis for each miRNA, so you need # miRNAs x # samples amount of reverse transcription reactions. It's a lot of reverse transcription reactions.

The second way is Qiagen's "miRCURY" method which I used to validate miRNA-seq in my 2021 first author preprint (see my profile). You only need one cDNA per sample and you can use it for several miRNAs. The random primers bind your miRNA pool and add a "5' universal tag" sequence during the reverse transcription step. All your cDNA has this universal tag, so it isn't miRNA-specific yet. The next step is to order plates from Qiagen that come with probes that are specific for your miRNAs. The qPCR reaction makes fluorescence signal when the miRNA-specific probe is in double stranded DNA with the primers, so that is how the qPCR machine measures increasing amplification of your specific miRNAs. The Qiagen method was much easier when doing it, but the Qiagen ordering was confusing (some pages said you need X,Y,Z, other pages said you need only X and Z, etc) so I recommend you call a Qiagen rep and get them to help you figure out your experiment design.

2/I don't know what you mean by special box. Count the rows and columns of the pipette tip box. Are there 8 rows and 12 columns? If so, you have an 8x12 box with the same layout as a 96-well plate. If it's different, you can still organize something to work for your layout. My main point is that you should pair your tip box and PCR plate layout so that you are certain which PCR well you just finished, and which well you need to do next. Pipette tips #1-8 should be for PCR wells #1-8, etc. If something happens and you need two pipette tips per well (for example if the first tip was deformed), get the extra tip from another backup box. Don't lose your order.

I have been doing PCR for many years, but I still have moments in the middle of pipetting a 96-well plate when I lose my place (e.g. if someone interrupts me or I start daydreaming). It really helps reduce anxiety/mistakes if I am able to look at the pipette tip box and see, ok, I am on sample #20. I don't need to worry about adding cDNA #20 to PCR reaction #21 by accident because I lost my place. I highly recommend using your pipette tip layout to keep track of your experiment, especially for huge experiments like this. If you only rely on checking off samples on paper, someone can interrupt you or you can forget to check a box.

3/I found this calculator for primer stocks to final primer concentrations: https://www.biosearchtech.com/support/tools/oligo-toolbox/oligo-dilution-calculator

I prepare my PCR protocols on Excel. I like it because I can type up the 1X reaction, then add a column to get the volumes for a 16X stock very easily by just dragging a formula. I don't need to worry about typos or old recipes because Excel does the math for me. I also like to draw out my 96-well plate layouts so I can visualize the experiment before I do it. It is nice for lab meeting, when I am at the bench, and later during data analysis.

P.S. All this is advice for mature miRNAs. If I am wrong and your experiment is looking at longer sequences (like the original long transcripts) then a regular qPCR is ok.